We have previously described use of a Balch homogenizer for the rapid, cost-effective and freeze-thaw independent homogenization of Caenorhabditis elegans (Bhaskaran, S., et al., 2011). This tool operates through shear forces generated by placing a ball bearing of known diameter within a confined chamber and then using two disposable syringes to manually advance worm samples past the micron sized gap that is formed between the chamber wall and the bearing. Using this tool we showed that worms of any developmental stage could be equally well homogenized after selecting an appropriately sized bearing to precisely control the wall-to-bearing clearance. Here we describe the use of the Balch Homogenizer for isolation of nuclei from C. elegans.

Since the primary goal of this study was to identify conditions that permitted rapid, freeze-thaw independent isolation of structurally-sound nuclei from any stage of worm development, our choice of buffers as well as centrifuge spin speeds to pellet nuclei were the same as those described in the embryonic and oocyte nuclei purification protocol of Mains and McGhee (1999). In their method, several approaches to disrupt worm samples were described, including use of a vintage motorized Stansted Cell Disruptor, a French Press, sonication and a mortar & pestle. Each method had its associated limitations, ranging from cost to sample heating. We established the following protocol for nuclei extraction which is rapid, does not require prior sample freezing, and can be applied equally to all stages of nematode development. Samples of nuclei collected using this protocol have been utilized in a number of downstream applications, including preparation of nuclear extracts for use in gel shift assays.

For the following protocol we utilized a mixed-stage population of SD1084 worms containing a sur-5::GFP reporter gene. Use of this strain allowed us to monitor the degree of disruption of the nuclear membrane at each step of the protocol, as SUR-5::GFP is soluble. We have previously published ball bearing clearances required to either fracture or homogenize synchronized populations of worms at any stage of larval development [refer to Table 1 in (Bhaskaran, S., et al., 2011)]. The following protocol can therefore be easily modified to accommodate worms of any size. Nuclei in C. elegans range from ~1.8 – 7μm (Chen, L. et al., 2013).

Resuspend the pellet in 2-4 volumes of NEB. Transfer to a microfuge tube, extract by gentle rotation 45 minutes at 4°

Centrifuge at 10,000 x g for 14 minutes at 4°C to pellet debris.

Carefully remove the supernatant and store in aliquots at -80°C (Note: This sample represents a crude nuclear extract positive for GFP (Fig. 1A). Surprisingly it also contains buoyant structures that stain positive with propidium iodide (Fig. 1B). These structures are chromatin skeletons (Fig. 1C) and they presumably avoid precipitation, in part because of the high glycerol content of the extraction buffer.)

Figures

Figure 1: Characterization of extraction fractions using immunofluorescence and western analysis. (A) α-GFP western blot of various nuclei extraction fractions obtained using SD1084 worms. (Monoclonal GFP antibody obtained from Antibodies Inc., cat# 73-131). The precise identity of the expressed SUR-5::NLS::GFP fusion protein is unknown, but its full length predicted size is 104 kDa. (Yochem, J., et al., 1998). (B) Fluorescent images of mounted extract from Step 11 showing propidium iodide (PI) positive structures and SUR-5::GFP staining. (C) Higher magnification image of PI-positive structures shown in (B). Very few structures are both PI-positive and sur-5::GFP positive (compare left and right arrows), presumably due to detergent extraction of sur-5::GFP, but their presence nonetheless underscores their nuclear origin.

In the absence of microfluidic rigs, most live imaging protocols utilize flat agarose pads along with anesthetics and/or microbeads. It is often difficult to manipulate the worms position in this set up and thus a higher number of worms need to be mounted to ensure a few showing an optimal position for imaging. This is problematic when trying to image nematodes with genotypes that are scarce within a strain, such as m+z- sterile or embryonic/larval lethal, because finding enough animals to mount can be challenging. In order to image worms of genotypes that are hard to find, we slightly modified a previously published protocol (Zhang, M. et al., 2008) to make agarose pads that allow us to maintain L4 or adult hermaphrodites and adult males straight during imaging for at least 2 hours. These pads also appear to reduce rolling of the worms during imaging.

Materials:
A 12-inch long-playing (LP) vinyl record, labeling tape, Pasteur pipette with bulb, agar (at the concentration in which you prefer to use, we use 4% agar), microscope slides, your preferred anesthetic (we use either levamisole or serotonin), a minutien pin (eyelash pick), coverslips, a petroleum jelly (we use Vaseline) filled syringe with a flat end needle.

Procedure:
1. Place strips of tape 1.5-2 inches apart perpendicular to the grooves on the vinyl record’s grooved surface, as shown in Figure 1.A.
2. Pipette a drop of melted agarose onto the vinyl record between the strips of tape, and immediately add a microscope slide on top of the drop to make a pad against the vinyl record.
3. After cooling, remove microscope slide from the vinyl record by lifting from one side as shown in Figure 1.B. If the pad is too large, it can be trimmed down by using another microscope slide’s flat edge to make straight cuts.
4. Add a volume of 2-5l of anesthetic at a time, depending on the size of your pad. Then pick worms onto anesthetic droplet (the grooves are an optimal width for hermaphrodites from L4-adult).
5. Position worms into the grooves in the agarose pad utilizing the minutien pin (Figure 1.C). Worms are easier to position when most of the anesthetic has been absorbed into the pad, but don’t allow it to dry completely, as this will form air bubbles in the adjacent grooves.
6. Make a circle line around the pad with the syringe filled with Vaseline before placing the coverslip as shown in Figure 1D. Slightly press the coverslip over the pad and vaseline to keep the worms in place and make a seal around the pad to prevent evaporation of the anesthetics mix.

This protocol can be used in conjunction with microbeads, but in our hands they do not seem to improve the immobilization of the worms.

The vinyl record grooved pads function, in part, as a poor-person’s microfluidics for live imaging. It allows the placing of worms into single channels organized as in a microfluidic chip, thus reducing the time searching for the worms for imaging on the slide. Addition of drugs or food (for prolonged imaging) could potentially be done with a fine needle inserted through the Vaseline ring. An additional advantage of these pads is that there is no difference in fluorescence brightness loss compared to the conventional agarose pad.

This protocol’s affordability, the ease of use and training has simplified our live imaging of m+z- males expressing low fluorescence fusion proteins in the germline significantly.

We thank the Mikaela Murph and Erlyana Clarke for trying the protocol and providing us with feedback.

Figures

Figure 1: Figure 1: Overview of the process for making agar pads using a vinyl record surface. A. Laboratory label tape placed on vinyl disc gives reference where to place the agar drop and also establishes an even pad thickness. B. A drop of hot agar is covered with the microscope slide. Once agar has cooled down slide is lifted with the attached agar pad. C. Anesthetized C. elegans nematode aligned on a groove made with vinyl record. D. Vaseline perimeter is applied around the agar pad and slide cover is pressed slightly to seal in the humidity to prevent the nematodes from drying over long periods of imaging.

One of the many advantage of working with C. elegans is the ability to make frozen stocks from which viable animals can be recovered as needed. While we often brag about this to our fly colleagues, we all know that the truth is that worms really don’t freeze all that well. When thawing an aliquot of frozen stock, we are happy if we can recover more than just a few viable larvae amongst the mass of dead worms. Hoping to develop a better method for freezing worms, I turned to the literature on freezing plant cells which are difficult to freeze as cryoprotectants are often unable to penetrate the thick cell walls. Since the cuticle of nematodes might also block the penetration of some cryoprotectants, I decided to try a cryoprotectant that works on plant cells.The following is a protocol that I have modified for use in worms and have employed for the past 5 years. I have found that it is consistently far superior to the traditional glycerol-based method and allows the recovery of a sizable recovery of the frozen worms including all larval stages and adults.

Freezing Worms with Trehalose-DMSO

Grow worms on 1-3 100 mm plates until freshly starved. Generally the more worms the better.

Wash worms off plates in M9 buffer and collect by centrifugation (5000 rpms for 3 minutes is sufficient.

We are pleased to invite student and postdoc abstract submissions for 5 minute lightning talks to be presented at the NSF-sponsored, 3rd Parasitic Nematodes: Bridging the Divide workshop. We encourage submissions that address the broad theme of C. elegans as a model for parasitic nematode biology, including work relating to human, animal, and plant parasitic nematodes and from both a basic biology standpoint or a translational perspective.

The platform provided by this workshop represents a unique opportunity for early-career scientists to interact with other researchers at the intersection of C. elegans biology and parasitology. These five minute talks will be presented on the first day of the 21st International C. elegans Meeting. The presenting authors must be available during any of the limited number of possible time slots available.

Abstract submissions are due on May 12th at 5pm PST and presenting authors will be notified of decisions by May 19th. Instructions for logistics and formatting of lightning talk presentations will follow acceptance. Abstract Format: Abstracts should be limited to 2,500 characters in total, including the title, authors, author affiliations, main body, and spaces. Abstracts should be sent to the workshop e-mail address (parasiteworkshop@gmail.com) as a single PDF file with subject heading “Lightning Talk Abstract”. A notification of receipt will follow submission. Accepted abstracts will appear online in a Worm Breeder’s Gazette article and titles will be listed on the International Worm Meeting website.

Reliable and sensitive tools for labeling specific connections between individual neurons are crucial for our understanding of the functionality of the nervous system. The gold standard for visualizing synaptic connections is electron microscopy (EM), wherein the degree of connectivity (number of sections over which en passant synapses are observed) and ultrastructure can be resolved. However, given the substantial time and resource investment, EM reconstruction is impractical for routine analysis of synaptic connections under distinct conditions (e.g. different time points, developmental stages, genetic backgrounds) and even more so for dynamic studies of neuronal connections. The genetically controlled GRASP system (“GFP Reconstitution Across Synaptic Partners”) has been used to visualize synaptic connections in live, transgenic animals (Feinberg et al., 2008). GRASP is based on two non-fluorescent split-GFP fragments, spGFP1-10 and spGFP11, tethered to synaptic membranes in each of two specific neurons or cell-types. When two neurons, each expressing one of the fragments, are tightly apposed across a synaptic cleft, fluorescent GFP is reconstituted. Actual synapses are labeled by targeting the Split GFP molecules to synaptic membranes using the neuroligin protein. C. elegans Neuroligin localizes both pre- and postsynaptically in neurons.

We have recently used GRASP to label a substantial number of synapses in C. elegans (Oren-Suissa et al., 2016)(E.A.B., M.H., M.M., A.B. and O.H., unpubl. data) and want to share a few lessons that we learned:

1) To best visualize the area of contact between the two cells of choice, the neuronal processes should be labeled with a fluorescent cytoplasmic marker. Using both cellular markers in the same color is an option as long as the area of contact is clearly visible. Optimally, promoters should differ from those driving the split GFP fragments. The cell-specificity is less of a problem and promoters for cytoplasmic labeling could be expressed in additional cells.

2) We perform germ line transformation by microinjection. In cases where the GRASP signal is expected in ventral or dorsal areas of the body (for example, the PAG), we prefer to use the pRF4 (rol-6(su1006)) plasmid as the co-injection marker. It allows for easy visualization of GRASP puncta, as the roller worm is slightly rotated when mounted on a slide for imaging purposes. We co-inject 5 plasmids, two split GFP plasmids, two cytoplasmic markers and a co-injection marker. Alternatively, the GRASP construct could be injected into an already integrated cytoplasmic marker strain, so the marker and GRASP do not need to be on the same array.

3) Initial split GFP concentrations should be minimal and aimed at 10 ng/mcL. If same promoters are used for the cytoplasmic markers, the latter should be injected in extremely low concentrations, to avoid competition for transcription resources. All the heritable F2 worms are maintained for initial live-imaging analysis. As many transgenic lines as possible should be screened to uncover ones with synaptic GFP puncta localization. Imaging should be carried out as described below. Positive transgenic lines are those with discrete GFP puncta between the two cells. If expression is too high, synapses are saturated with GFP and the split-GFP interaction is irreversible. Transgenic animals generated by microinjection carry large extrachromosomal arrays incorporating a high copy number of transgenes, which might cause overexpression and germ line silencing. In addition, the inherit­ance of extrachromosomal arrays is unstable and can create genetic mosaics. We integrate the extrachromosomal arrays into C. elegans chromosomes for stable inheritance, using γ-irradiation. In some cases integrated GRASP strains underwent silencing. Growing these strains at 25C for a couple generations disilenced these arrays.

4) Transgenics lines with good signals are often hard to find. Concentration of split-GFP plasmids should be gradually increased, either of both plasmids or of one of them. A common solution is to swap the split GFP fragment between the pre- and the post-synaptic cells. Alternatively, different neuronal promoters should be tried.

5) GRASP imaging conditions: Compared to our Zeiss Imager Z1 standard compound epifluorescence microscope, we have observed much better signals on a pinhole based confocal microscope (LSM880). Epi-fluorescent microscope images contain too much out-of-focus signals. Puncta were quantified by scanning the original full Z-stack for distinct dots in the area where the processes of the two neurons overlap.

Share this:

Like this:

About The WBG

The Worm Breeder's Gazette publishes brief articles on new findings, experimental techniques, meetings and other things of interest to the C. elegans and nematode research communities. Read more about The WBG, what we publish, and how to participate.